Aphis gossypii (cotton aphid)
- Summary of Invasiveness
- Taxonomic Tree
- Notes on Taxonomy and Nomenclature
- Distribution Table
- History of Introduction and Spread
- Habitat List
- Hosts/Species Affected
- Host Plants and Other Plants Affected
- Growth Stages
- List of Symptoms/Signs
- Species Vectored
- Biology and Ecology
- Natural enemies
- Notes on Natural Enemies
- Means of Movement and Dispersal
- Plant Trade
- Impact Summary
- Environmental Impact
- Impact: Biodiversity
- Social Impact
- Detection and Inspection
- Similarities to Other Species/Conditions
- Prevention and Control
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- Aphis gossypii Glover, 1877
Preferred Common Name
- cotton aphid
Other Scientific Names
- Aphis bauhiniae Theobald, 1918
- Aphis circezandis Fitch, 1870
- Aphis citri Ashmead of Essig, 1909
- Aphis citrulli Ashmead, 1882
- Aphis cucumeris Forbes, 1883
- Aphis cucurbiti Buckton, 1879
- Aphis helianthi Monell
- Aphis heraclella
- Aphis heraclii
- Aphis lilicola Williams, 1911
- Aphis minuta Wilson, 1911
- Aphis monardae Oestlund, 1887
- Aphis parvus Theobald, 1915
- Aphis tectonae van der Goot, 1917
- Cerosipha gossypii (Glover, 1877)
- Doralina frangulae (Kaltenbach)
- Doralina gossypii (Glover)
- Doralis frangulae (Kaltenbach)
- Doralis gossypii (Glover, 1877)
- Toxoptera leonuri Takahashi, 1921
International Common Names
- English: betelvine aphid; cucurbits aphid; green aphis; melon aphid
- Spanish: afido del algodón; afido del melón; piojo del algodon; pulgón del algodonero; pulgón del melón
- French: puceron du cotonnier; puceron du melon
- Portuguese: pulgao das cucurbitaceas; pulgao do algodoeiro
Local Common Names
- Denmark: agurkbladlus
- Germany: gruene baumwoll-blattlaus; gruene gurken-blattlaus; melonen-blattlaus
- India: brinjal aphid
- Iran: schatte khiar (sabs rang)
- Israel: knimat hadeluyim
- Japan: wata-aburamusi
- Mexico: pulgon del algodonero o mielecilla
- Netherlands: groene katoenbladluis; katoenluis
- Turkey: pamuk yaprak biti
- APHIGO (Aphis gossypii)
Summary of InvasivenessTop of page
This aphid is not in the Invasive Global Species Database as an invasive pest (ISSG, 2006). However, it is registered in the Invasive species database of the United States Department of Agriculture (Anon., 2005). It is unclear if entry into this database indicates an official classification of this aphid as 'invasive', or if its presence is entirely because it is not native to North America.
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Metazoa
- Phylum: Arthropoda
- Subphylum: Uniramia
- Class: Insecta
- Order: Hemiptera
- Suborder: Sternorrhyncha
- Unknown: Aphidoidea
- Family: Aphididae
- Genus: Aphis
- Species: Aphis gossypii
Notes on Taxonomy and NomenclatureTop of page
There have been several biotype designations used for A. gossypii. However, it should be treated as a single exceptionally variable species, until meaningful and standardized quantification of the diversity within the species can be undertaken. Although A. gossypii has been reported from many hosts, individual clones do not have the ability to feed on all reported hosts (Liu et al., 2002).
Taxonomically, A. gossypii is difficult to separate from Aphis frangulae. Strogani (1984) summarizes information about the A. frangulae group available at that time. He regarded gossypii as a sub-species of A. frangulae.
DescriptionTop of page
The cornicles or siphunculi are uniformly sclerotized from tip to base, and darkly pigmented. They are longer than the cauda and gradually taper towards the apex with a small dilation there.
The dorsal abdominal segments are uniformly sclerotized and unpigmented. A. gossypii reared near the colour transition temperature (see later in this section) often have a blotchy appearance with some parts greenish and other parts more yellow. The cauda usually have 4-7 hairs and are paler than the cornicles. The metafemoral hairs are all shorter than the basal width of the metafemur. Stidulatory apparatus is absent. The antennal tubercles are weakly developed. The terminal process is more than twice the length of the last antennal segment, but less than 3.5 times as long. The rostrum has a blunt apex and the terminal rostral segment shorter than cauda and under 1.5 times the length of the second metatarsal segment.
A. gossypii is a small aphid. Adults range from just under 1-1.5 mm in body length. The minimum diameter is just over 0.34 mm. This is indicated by the fact that a screen with a mesh diameter of under 0.34 mm is able to exclude A. gossypii (Bethke and Paine, 1990).
Cornicle length distinguishes separate instars of A. gossypii reared under fluctuating temperatures. There is considerable overlap between instars, but none between nymphs and adults (Singh and Srivastava, 1989). Other characteristics combined can provide greater separation between instars especially under constant temperatures (Inaizumi and Takahashi, 1989a; Ebert and Cartwright, 1997).
First-instar nymphs are distinguished by having 4 antennal segments, whereas second-instar nymphs have 5 segments. Differences between second- and third-instar nymphs are fairly small, but at constant temperatures they can be distinguished using a combination of characteristics. Third-instar nymphs have no setae on the margin of the genital plate, whereas fourth-instar nymphs have such setae. Second-instar nymphs with developing wings appear to have shoulders, third-instar nymphs have small wing pads, and the developing wings are prominent on fourth-instar nymphs (Ebert and Cartwright, 1997).
The occurrence of the following A. gossypii stages have been described in Japan: fundatrix, fundatrigeniae, alienicola, gynoparae, oviparous female, alate and apterous male, hibernating viviparae, virgin androparae, androparae, heteroparae and androgynoparae. There are also individuals with partly developed wings from nearly apterous to nearly functionally winged (Inaizumi, 1968; Inaizumi, 1980; Inaizumi, 1983). Miyazaki (1987), Moran (1992) and Blackman (1994) have defined the life stages of A. gossypii. The following stages have been described for A. gossypii in Iran: alate and apterous viviparae (Ghovanlou, 1974; Ebert and Cartwright, 1997).
A. gossypii can range in colour from yellow to very dark (almost black) green. The smaller yellow form occurs during warmer summer conditions. The green form is larger and occurs during cooler spring and autumn temperatures, and uncrowded conditions. Colour morphs are able to produce progeny of another colour morph (Wall, 1933; Setokuchi, 1981). Host plant can also influence aphid colour (Regupathy and Jayaraj, 1973). Wall (1933) reported that green morphs produce more alate offspring than do yellow morphs. However, his observation could be the result of crowding, as the green form also produced more total offspring (Ebert and Cartwright, 1997).
The use of a proper key is highly recommended and the aphids should be examined by an expert for confirmation.
DistributionTop of page
Distribution TableTop of page
The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.Last updated: 25 Feb 2021
History of Introduction and SpreadTop of page
Habitat ListTop of page
Hosts/Species AffectedTop of page
The host range of A. gossypii (over 92 plant families) includes food and fibre crops, ornamentals and flowers. Ebert and Cartwright (1997) have compiled a list of hosts by plant family, and a list of all plant families that contain at least one potential host. An exhaustive list of plant species which serve as hosts in Australia and the Pacific Ocean can be found in Carver (1996).
The host list in this Compendium is a list of crops where A. gossypii is able to feed and reproduce. The list of non-crop plants that can serve as hosts is more extensive. A more comprehensive list can be found in the references cited in Ebert and Cartwright (1997). Primary host crops are ones where the aphid can cause plant mortality through direct damage and vectors one or more viruses worldwide. Secondary host crops are ones where the aphid is not a global problem, or where it either does not cause much direct damage or is not a vector.
Host Plants and Other Plants AffectedTop of page
Growth StagesTop of page
SymptomsTop of page
A. gossypii will attack most parts of the plant if population density is high enough. Exceptions include direct feeding on mature reproductive structures (fruits, berries, nuts) and feeding on roots. However, even though there is no direct damage to these structures, a general decline in plant health will affect the proper development of these tissues.
List of Symptoms/SignsTop of page
|Growing point / distortion|
|Inflorescence / honeydew or sooty mould|
|Inflorescence / twisting and distortion|
|Leaves / abnormal colours|
|Leaves / abnormal forms|
|Leaves / honeydew or sooty mould|
|Leaves / honeydew or sooty mould|
|Leaves / honeydew or sooty mould|
|Leaves / wilting|
|Stems / stunting or rosetting|
|Whole plant / dwarfing|
|Whole plant / external feeding|
Species VectoredTop of page
Bean common mosaic necrosis virus
Bean common mosaic virus (common mosaic of beans)
Bean leafroll virus (pea leafroll virus)
Bean yellow mosaic virus (bean yellow mosaic)
Beet yellows virus (beet yellows)
Carnation mottle virus (mottle of carnation)
Cauliflower mosaic virus (cauliflower mosaic)
Citrus tristeza virus (citrus tristeza)
Cowpea aphid-borne mosaic virus
Cucumber mosaic virus (cucumber mosaic)
Cucurbit aphid-borne yellows virus (Cucurbit aphid-borne yellows)
East Asian Passiflora virus
Lettuce mosaic virus (lettuce mosaic)
Lily symptomless virus
Maize dwarf mosaic virus (dwarf mosaic of maize)
Onion yellow dwarf virus (onion yellow dwarf)
Papaya ringspot virus
Passion fruit woodiness virus (passionfruit woodiness disease)
Peanut mottle virus (peanut mottle)
Peanut stripe virus (groundnut stripe disease)
Pepper veinal mottle virus
Plum pox virus (sharka)
Potato leafroll virus
Potato virus Y (potato mottle)
Sugarcane mosaic virus (sugarcane mosaic)
Sweet potato feathery mottle virus (internal cork disease of sweet potato)
Tobacco etch virus (tobacco etch)
Tulip breaking virus
Turnip mosaic virus (cabbage A virus mosaic)
Watermelon mosaic virus (watermelon mosaic)
Watermelon mosaic virus 1 (WMV - strain 1)
Yam mosaic virus
Zucchini yellow mosaic virus
Biology and EcologyTop of page
A. gossypii reproduction is mostly asexual with either alate or apterous females. In warmer environments, A. gossypii exhibits an anholocyclic life cycle, while in cooler areas it exhibits either a heteroecious or autoecious holocyclic life cycle (Slosser et al., 1989; Zhang and Zhong, 1990). The heteroecious cycle involves a migration from a winter host to a summer host in the spring and a return to a winter host in the autumn for laying eggs.
Reproductive rates in A. gossypii are reported in two ways; net reproductive rate which is an interaction between birth rate and survival rate (Wilson and Bossert, 1971), and birth rate as measured in nymphs produced per day per aphid.
Net reproductive rate has been estimated for A. gossypii on many hosts: squashes (Cucurbita pepo) (Aldyhim and Khalil, 1993), citrus (Citrus unshiu) (Komazaki, 1982), pumpkins (Liu and Perng, 1987), Veronica persica (Nozato, 1987a), taro (Setokuchi, 1981) and cucumbers (Wyatt and Brown, 1977; Kocourek et al., 1994; Owusu et al., 1994b; van Steenis and El-Khawass, 1995).
Significant differences were found in birth rates of A. gossypii reared on cotton, watermelons and groundnuts (Ekukole, 1990). Significant differences were also found in its reproductive potential on cotton, watermelons, sesame and aubergines (Moursi et al., 1985). Another study reported that development time was shortest on cotton, longest on melons, with development rate on watermelons lying somewhere between (Ghovanlou, 1976). In contrast, one paper reported no significant differences in birth rate for A. gossypii reared on okra, aubergines and chillis (Kandoria and Jamwal, 1988).
A. gossypii takes 5.2 days to reach maturity on cotton at 28°C. The optimal temperature for reproduction is 20-25°C when the aphid can produce an average of 2.8 nymphs per day (in the USA) (Isely, 1946; Akey and Butler, 1993).
There are two forces proposed as triggers for alate production in A. gossypii: nutritional factors and crowding. However, research to date has not conclusively identified the relative importance of nutrition versus crowding in alate formation in A. gossypii. Nutritional stress has been examined by removing aphids from their host for some period of time and then returning them to their host. This treatment was applied to adults and nymphs on cotton for 6-8-hour periods each day. Starved nymphs from apterous parents resulted in 13% alates versus 0.4% from unstarved nymphs. However, starvation of nymphs from alate parents resulted in no increase in alate formation. A similar result was also reported for starved parents, where starved apterous parents produced more alate progeny than well-fed apterous parents (23% versus 2%, respectively) and there was no increase in alate formation by starved alate parents. The effect of starvation was also detectable in progeny from aphids starved as nymphs but fed normally as adults (Reinhard, 1927). However, this might be a crowding response if A. gossypii interprets physical contact with the brush (used to remove the aphid from the plant) as contact with other aphids.
Nutritional factors from other sources can affect A. gossypii development. In considering the possible role of aphid-borne plant viruses, the survival of the virus is dependent on having an efficient aphid vector, and the most efficient vector is alate. Therefore one might expect that a virus would promote conditions favouring alate production in the aphid. Alate production in the melon aphid-courgette-Zucchini yellow mosaic virus system appeared to increase on infected plants. Unfortunately, virus-infected plants also had more aphids. Blua (1991) and Blua and Perring (1992a) decided that nutritional factors are the cause for increased alate production. To further support the argument that the cause might be nutritional, it has been shown that aphid infestations on Solanum integrifolium changes peroxidase, esterase and protein content of the plant in proportion to the level of infestation (Owusu et al., 1994a). This shows a change in plant nutrient content associated with aphid density which might form a chemical link between nutrition and alate production. More work in this area is required.
Crowding effects have been shown for A. gossypii on cotton. One study examined crowding using leaf disks with a single apterous aphid which was removed following reproduction. The resulting colonies contained 1-7 nymphs. From 52 colonies with 1-2 nymphs, no alates were produced. From 41 colonies with 3-4 nymphs, less than 10% of the colony became alate. However, of 29 colonies with 5-7 nymphs, over 30% of the total number of aphids became alate (Graham, 1968). This experiment was repeated as part of the control for another experiment. Two possibly significant differences were apparent: alate production only occurred in colonies with four or more nymphs, and colonies with 5-7 nymphs only produced 12% alates (Graham, 1968). The observation that solitary individuals never develop alates was reported earlier for the melon aphid (Reinhard, 1927).
Additional factors which influence alate production include crowding of parents (Graham, 1968), the type of parent (apterous or alate) (Reinhard, 1927), and the possible effects of light and temperature (Reinhard, 1927; Guldemond et al., 1994).
Temperature is one of the most important abiotic factors affecting the life cycle of A. gossypii. A lower developmental threshold for A. gossypii was estimated at 7.34°C on squash in Taiwan (Liu and Perng, 1987). Development thresholds have also been estimated for A. gossypii on cucumbers as 5.8°C from birth to age of first reproduction, but the development threshold for the nymphal stages was 6.9°C (Kocourek et al., 1994). A study of A. gossypii on Veronica persica estimated a developmental threshold of 10.47°C for the teneral preflight period (Nozato, 1989b). An upper limit to A. gossypii survival of 35°C was reported on squashes in Saudi Arabia, but it also survives in okra fields where the daytime temperature exceeds 45°C (Aldyhim and Khalil, 1993). Temperature is also thought to be responsible for some strains of A. gossypii being holocyclic whereas others are anholocyclic. A hypothesis that eggs will be produced in locations where the average temperature during November does not exceed 13°C has been proposed (Inaizumi, 1980).
Light intensity and day length are also important in the reproductive capacity of A. gossypii. Increasing daylength increases reproductive rate (Aldyhim and Khalil, 1993). The effect of light intensity is unclear, and higher intensity increased reproduction between 800 and 8000 Lux (Wyatt and Brown, 1977), while it decreased reproduction between 54 and 550 Lux (Auclair, 1967).
Flight is the beginning of the dispersal phase in the A. gossypii life cycle. It begins with the preflight period (from moult to flight) which lasted from 1 to 31 hours with most activity 10-24 hours after moult from colonies reared on Veronica persica (Nozato, 1987b). The teneral preflight period increased from 10 to over 70 hours with decreasing temperatures from 28 to 12°C (Nozato, 1989b). Adults flew from about sunrise to early afternoon, but a few individuals continued to fly after dark. With first light at 06.00 h, and last light at 19.30 h, no flight was detected from 23.00 h to 07.00 h. Considering that the time of moulting is independent of time of day, the most common duration of the teneral preflight period is explained by adult inactivity after dark (Nozato, 1987b, 1989b).
In laboratory colonies, the flight period lasted from 1 to 4 days (Nozato, 1990). Older colonies produced fewer alates that flew for 1 day and more that flew for 2 days. Aphids flew from one to several (about 5) times each day, with the first flight always longer than the others. Alates larviposited after flight, and flew again when the number of embryos with pigmented eyes per ovariole decreased. Alates that flew longer had a shorter reproductive period and produced fewer total progeny (Nozato, 1987b). Reports indicate that alates will not produce offspring on leaves with existing colonies (Reinhard, 1927; Nozato, 1989a).
In Côte d'Ivoire, dispersal from savannah to cotton fields was examined for A. gossypii from data collected using pan traps. The data showed that most aphids settled at field margins, although there was some settlement in the field. Pan traps at ground level caught more A. gossypii than traps further from the ground and most aphids were caught no more than 1 m from the soil surface (the closer to the surface, the more aphids were caught) (Duviard et al., 1976).
Orientation to host plants was significant at 6 hours after wing development, but was highly significant after 24 hours. Alates were able to distinguish between different plants; Cucurbita pepo and Thunbergia laurifolia were attractive, and were common hosts for A. gossypii in Cuba. The occasional host Hibiscus rosa-sinensis was neither attractive nor repellent, and the non-host plant Lantana camara was repellent (Pospisil, 1972). (L. camara found in locations other than Cuba has been recorded as a summer host of A. gossypii. This apparent contradiction is due to differences in aphid populations found in differing geographical areas.)
Egg laying on H. syriacus occurred mostly between the leaf scar and the twig near where the buds would emerge in spring. Some eggs were also laid at the branching point of twigs. However, from the wandering behaviour of the oviparous A. gossypii females, it appears that they searched for protected places to lay eggs rather than for specific parts of the plant (Inaizumi and Takahashi, 1989b).
The most important impact A. gossypii has on world agriculture is through its ability to transmit plant viruses. See Kennedy et al. (1962) and Ebert and Cartwright (1997).
A. gossypii is a vector for the following viruses:
Alfalfa mosaic virus
Bean common mosaic virus
Calotropis ringspot mosaic virus
Carnation mottle virus
Cauliflower mosaic virus
Chinese yam necrotic mosaic virus
Citrus tristeza virus
Citrus woody gall vrus
Commelina mosaic virus
Cowpea (aphid-borne) mosaic virus
Cucumber mosaic virus
Garlic mosaic virus
Greengram mosaic virus
Infectious chlorosis of banana
Leaf crinkle of sunflower
Lily symptomless virus
Muskmelon yellow stunt virus
Onion yellow dwarf virus
Papaya ringspot virus
Passionfruit Sri Lankan mottle virus
Pepper veinal mottle virus
Potato leafroll virus
Potato virus Y
Solanum trovum mosaic virus
Sri Lankan passionfruit mottle virus
Sugarcane mosaic virus
Sweet potato feathery mottle virus
Turnip mosaic virus
Watermelon mosaic 1 virus
Watermelon mosaic 2 virus
Yam mosaic virus
Sunflower yellow blotch virus
Yellow vein mosaic virus
Zucchini yellow mosaic virus
A. gossypii on okra infected with Yellow-vein mosaic virus did not reproduce as fast as those on healthy okra (Regupathy and Jayaraj, 1972). A. gossypii on courgette plants infected with Zucchini yellow mosaic virus spent more time probing and less time feeding than those on healthy plants. Furthermore, prior to feeding, aphids on infected plants spent more time in forming the salivary sheath (Blua and Perring, 1992b).
Natural enemiesTop of page
|Natural enemy||Type||Life stages||Specificity||References||Biological control in||Biological control on|
|Adalia bipunctata||Predator||Adults; Arthropods|Nymphs||Azores; USSR|
|Allograpta exotica||Predator||Sturza et al. (2011)|
|Alloxysta pleuralis||Parasite||Ferrer-Suay et al. (2014)|
|Amblyseius swirskii||Predator||Adults; Arthropods|Nymphs|
|Anisosticta novemdecimpunctata||Predator||Adults; Arthropods|Nymphs|
|Anystis baccarum||Predator||Adults; Arthropods|Nymphs|
|Aphelinus abdominalis||Parasite||Adults; Arthropods|Nymphs|
|Aphelinus gossypii||Parasite||Adults; Eggs; Arthropods|Larvae; Arthropods|Nymphs; Arthropods|Pupae|
|Aphelinus mali||Parasite||Adults; Arthropods|Nymphs|
|Aphelinus paramali||Parasite||Adults; Arthropods|Nymphs|
|Aphelinus semiflavus||Parasite||Adults; Arthropods|Nymphs|
|Aphelinus varipes||Parasite||Adults; Arthropods|Nymphs||California; Paraguay||cereals; cotton|
|Aphidius colemani||Parasite||Adults; Arthropods|Nymphs||Netherlands|
|Aphidius gifuensis||Parasite||Adults; Arthropods|Nymphs|
|Aphidius matricariae||Parasite||Adults; Arthropods|Nymphs||Germany|
|Aphidius picipes||Parasite||Adults; Arthropods|Nymphs|
|Aphidius uzbekistanicus||Parasite||Adults; Arthropods|Nymphs|
|Aphidoletes aphidimyza||Predator||Adults; Arthropods|Nymphs|
|Beauveria bassiana||Pathogen||Adults; Arthropods|Nymphs|
|Binodoxys communis||Parasite||Adults; Arthropods|Nymphs||Taiwan||aubergines|
|Brinckochrysa scelestes||Predator||Adults; Arthropods|Nymphs|
|Callicaria superba||Predator||Adults; Arthropods|Nymphs||India; Manipur||Chromolaena odorata|
|Calotes versicolor||Predator||Adults; Arthropods|Nymphs|
|Cheilomenes lunata||Predator||Adults; Arthropods|Nymphs|
|Cheilomenes propinqua||Predator||Adults; Arthropods|Nymphs|
|Cheilomenes sexmaculata||Predator||Adults; Arthropods|Nymphs||Karnataka; Manipur; Taiwan|
|Cheilomenes sulphurea||Predator||Adults; Arthropods|Nymphs|
|Chrysemosa jeanneli||Predator||Youssif et al. (2014)|
|Chrysopa formosa||Predator||Adults; Arthropods|Nymphs||China; Shandong||cotton|
|Chrysopa intima||Predator||Adults; Arthropods|Nymphs||China; Shandong||cotton|
|Chrysopa orestes||Predator||Adults; Arthropods|Nymphs|
|Chrysopa pallens||Predator||Adults; Arthropods|Nymphs||Shandong|
|Chrysoperla carnea||Predator||Adults; Arthropods|Nymphs||Russia; USSR|
|Chrysoperla rufilabris||Predator||Adults; Arthropods|Nymphs|
|Chrysoperla sinica||Predator||Adults; Arthropods|Nymphs||Shandong||cotton|
|Coccinella septempunctata||Predator||Adults; Arthropods|Nymphs||China; China; Shandong; Honan; India; Manipur; Moldova; Russian Far East; USSR||Ageratum conyzoides; cotton|
|Coccinella septempunctata brucki||Predator||Adults; Arthropods|Nymphs|
|Coccinella transversalis||Predator||Adults; Arthropods|Nymphs||India; Manipur||Ageratum conyzoides|
|Coccinella undecimpunctata||Predator||Adults; Arthropods|Nymphs||Turkmenistan||cotton|
|Coelophora saucia||Predator||Adults; Arthropods|Nymphs||Manipur; Taiwan|
|Coleosoma octomaculatum||Predator||Adults; Arthropods|Nymphs||China; Shandong||cotton|
|Conidiobolus obscurus||Pathogen||Adults; Arthropods|Nymphs|
|Conidiobolus thromboides||Pathogen||Adults; Arthropods|Nymphs|
|Cryptogonus horishanus||Predator||Adults; Arthropods|Nymphs||Taiwan||aubergines|
|Cycloneda ancoralis||Predator||Adults; Arthropods|Nymphs|
|Cycloneda limbifer||Predator||Adults; Arthropods|Nymphs||USSR|
|Cycloneda sanguinea||Predator||Adults; Arthropods|Nymphs|
|Deraeocoris nebulosus||Predator||Adults; Arthropods|Nymphs|
|Deraeocoris pallens||Predator||Adults; Arthropods|Nymphs|
|Deraeocoris signatus||Predator||Adults; Arthropods|Nymphs|
|Diaeretiella rapae||Parasite||Adults; Arthropods|Nymphs|
|Ephedrus cerasicola||Parasite||Adults; Arthropods|Nymphs|
|Ephedrus persicae||Parasite||Adults; Arthropods|Nymphs|
|Ephedrus plagiator||Parasite||Adults; Arthropods|Nymphs|
|Episyrphus balteatus||Predator||Adults; Arthropods|Nymphs|
|Erigonidium graminicolum||Predator||Adults; Arthropods|Nymphs||China; Shandong||cotton|
|Euborellia pallipes||Predator||Adults; Arthropods|Nymphs|
|Eupeodes corollae||Predator||Adults; Arthropods|Nymphs||USSR; Sardinia||cucumbers|
|Geocoris jucundus||Predator||Adults; Arthropods|Nymphs|
|Gyrocaria sauzeti||India; Manipur||Chromolaena odorata|
|Harmonia axyridis||Predator||Adults; Arthropods|Nymphs||China; China; Shandong; Russian Far East; USSR||cotton|
|Harmonia conformis||Predator||Adults; Arthropods|Nymphs|
|Harmonia dimidiata||Predator||Adults; Arthropods|Nymphs||India; Manipur||Sesamum|
|Harmonia octomaculata||Predator||Adults; Arthropods|Nymphs||India; Manipur||Luffa aegyptiaca|
|Harmonia oxyridis||Predator||Adults; Arthropods|Nymphs||Azores|
|Hippodamia convergens||Predator||Adults; Arthropods|Nymphs|
|Hippodamia variegata||Predator||Adults; Arthropods|Nymphs||China; China; Shandong; Turkmenistan||cotton|
|Hyperaspis senegalensis||Predator||Adults; Arthropods|Nymphs|
|Ischiodon aegyptius||Predator||Adults; Arthropods|Nymphs|
|Ischiodon scutellaris||Predator||Adults; Arthropods|Nymphs||India||Cucumis|
|Lecanicillium lecanii||Pathogen||Adults; Arthropods|Nymphs||Netherlands; UK||chrysanthemums|
|Lipolexis gracilis||Parasite||Adults; Arthropods|Nymphs||India; Jammu and Kashmir||Zinnia elegans|
|Lipolexis pseudoscutellaris||Parasite||Adults; Arthropods|Nymphs|
|Lipolexis scutellaris||Parasite||Adults; Arthropods|Nymphs|
|Lysiphlebia japonica||Parasite||Adults; Arthropods|Nymphs|
|Lysiphlebia mirzai||Parasite||Adults; Arthropods|Nymphs|
|Lysiphlebus confusus||Parasite||Adults; Arthropods|Nymphs||Yunnan|
|Lysiphlebus delhiensis||Parasite||Adults; Arthropods|Nymphs|
|Lysiphlebus fabarum||Parasite||Adults; Arthropods|Nymphs||Morocco||Citrus; Pistacia|
|Lysiphlebus testaceipes||Parasite||Adults; Arthropods|Nymphs||France; Guadeloupe; Honduras; Ukraine||Ageratum conyzoides; Citrus; Cucurbita pepo; Hibiscus rosa-sinensis; watermelons|
|Mallada astur||Predator||Adults; Arthropods|Nymphs|
|Mallada basalis||Lu and Wang (2006)|
|Mallada boninensis||Predator||Adults; Arthropods|Nymphs|
|Micraspis discolor||Predator||Adults; Arthropods|Nymphs|
|Micromus posticus||Predator||Adults; Arthropods|Nymphs|
|Micromus timidus||Basu and Patro (2007)|
|Misumenops tricuspidatus||Predator||Adults; Arthropods|Nymphs||China; Shandong||cotton|
|Neda patula||Predator||Adults; Arthropods|Nymphs|
|Neozygites fresenii||Pathogen||Adults; Eggs; Arthropods|Larvae; Arthropods|Nymphs; Arthropods|Pupae|
|Nesidiocoris caesar||Predator||Adults; Arthropods|Nymphs|
|Ocyptamus gastrostactus||Predator||Adults; Arthropods|Nymphs|
|Oenopia kirbyi||Predator||Adults; Arthropods|Nymphs||India; Manipur||Duranta repens|
|Oenopia luteopustulata||Predator||Adults; Arthropods|Nymphs||India; Manipur||Chromolaena odorata; Duranta repens; Gynura lycopersifolia|
|Oenopia quadripunctata||Predator||Adults; Arthropods|Nymphs||India; Manipur||Chromolaena odorata; Duranta repens|
|Orius albidipennis||Predator||Adults; Arthropods|Nymphs|
|Orius minutus||Predator||Adults; Arthropods|Nymphs|
|Orius sauteri||Predator||Adults; Arthropods|Nymphs|
|Orius similis||Predator||Adults; Arthropods|Nymphs|
|Paecilomyces fumosoroseus||Pathogen||Adults; Arthropods|Nymphs|
|Paederus alfierii||Predator||Adults; Arthropods|Nymphs|
|Pardosa astrigera||Predator||Adults; Arthropods|Nymphs||China; Shandong||cotton|
|Pharoscymnus madagassus||Predator||Adults; Arthropods|Nymphs|
|Podisus maculiventris||Predator||Adults; Arthropods|Nymphs|
|Praon volucre||Parasite||Adults; Arthropods|Nymphs|
|Propylea japonica||Predator||Adults; Arthropods|Nymphs||China; China; Hubei; China; Shandong; Russian Far East; Taiwan||aubergines; cotton|
|Propylea quatuordecimpunctata||Predator||Adults; Arthropods|Nymphs||Moldova; Russian Far East; USSR|
|Pseudodoros clavatus||Predator||Adults; Arthropods|Nymphs|
|Pseudospidimerus circumflexus||Basu and Patro (2007)|
|Rhyzobius litura||Predator||Adults; Arthropods|Nymphs|
|Scymnodes lividigaster||Predator||Adults; Arthropods|Nymphs|
|Scymnus hoffmanni||Predator||Adults; Arthropods|Nymphs||Taiwan||aubergines|
|Scymnus interruptus||Predator||Adults; Arthropods|Nymphs|
|Scymnus louisianae||Predator||Adults; Arthropods|Nymphs|
|Scymnus pupulus||Predator||Adults; Arthropods|Nymphs|
|Sphaerophoria rueppellii||Predator||Adults; Arthropods|Nymphs||USSR|
|Sphaerophoria scripta||Predator||Adults; Arthropods|Nymphs||USSR||cucumbers|
|Spilocaria bissellata||Predator||Adults; Arthropods|Nymphs||India; Manipur||Ageratum conyzoides; Gynura lycopersifolia|
|Synonycha grandis||Predator||Adults; Arthropods|Nymphs||India; Manipur||okras|
|Tenodera sinensis||Predator||Adults; Arthropods|Nymphs|
|Trioxys angelicae||Parasite||Adults; Arthropods|Nymphs|
|Trioxys basicurvus||Parasite||Adults; Arthropods|Nymphs|
|Trioxys equatus||Parasite||Adults; Arthropods|Nymphs||India; Meghalaya||Urtica|
|Trioxys indicus||Parasite||Adults; Arthropods|Nymphs|
|Trioxys rubicola||Parasite||Adults; Arthropods|Nymphs|
|Trioxys sinensis||Parasite||Adults; Arthropods|Nymphs|
|Xysticus croceus||Predator||Adults; Arthropods|Nymphs||China; Shandong||cotton|
|Zelus renardii||Predator||Adults; Arthropods|Nymphs|
Notes on Natural EnemiesTop of page
The 'problem' with many predators is that they may only lay their eggs in the presence of sufficient aphids to ensure the survival of the larvae to maturity, and by then the damage may have been done. Non-specific predators with good searching ability that are active early in the year, such as Carabidae and Staphylinidae, are important when aphid numbers are low.
The predators of A. gossypii include a number of species in the Coccinellidae, Syrphidae, Chrysopidae, Hemerobiidae, and many small spiders. Although reports suggests that the predators will attack all life stages, there is no record of predators attacking the egg (Ebert and Cartwright, 1997).
The effectiveness of predators is highly variable, depending on the availability of alternative prey, host plant and environmental factors. The effect of alternative prey was reported for Chrysoperla rufilabris which preferred Heliothis virescens larvae to aphids, but preferred aphids to H. virescens eggs (Nordlund and Morrison, 1990). The presence of A. gossypii was shown to decrease predation on H. virescens eggs by Hippodamnia convergens, Chrysoperla carnea and Orius insidiosus (Ables et al., 1978).
The lacewing, Chrysoperla carnea, was able to cause an overall reduction in aphid abundance when caged on field-grown cotton in California, USA. Added to this system were several hemipteran generalist predators (Geocoris spp., Nabis spp. and Zelus spp.) which feed on A. gossypii, lacewing, and each other. All of these predators reduced aphid populations, though none were as effective as C. carnea. However, all of the hemipterans also reduced lacewing survival which resulted in an increased number of aphids. Furthermore, the reduction in the ability of lacewings to control aphid populations was increased with increasing size of the other predators (Rosenheim et al., 1993).
Syrphid flies have shown potential in controlling aphid populations under greenhouse conditions (Babayan and Hovhannisian, 1984; Chambers, 1986; Adashkevich and Karelin, 1988). However, colonization by the syrphid was decreased on older plants, and older larvae would not transfer from young plants to more mature plants (Adashkevich and Karelin, 1988). The suggested cause for the latter effect was leaf pubescence.
The most common parasites of A. gossypii are insects in the Hymenoptera. Additional parasites include members of the Diptera and some mites. While these parasites are able to parasitize all life stages except eggs, they prefer later instars (Ebert and Cartwright, 1997).
Changes in parasitism based on age structure of A. gossypii populations feeding on cotton were reported for the parasites Trioxys spp. and Aphelinus spp; these parasites rarely parasitized first- and second-instar aphids. Thus, the percentage of parasitism increased as the proportion of older aphids increased (Luo and Gan, 1986). This has survival value for both the parasite and A. gossypii because aphids which are parasitized as older nymphs or as adults have a chance to reproduce.
Aphids parasitized by Aphidius colemani had a fecundity of 0.5-1.3 nymphs/female when parasitized in the fourth-instar, and 10.5-13.3 when parasitized as adults (van Steenis and El-Khawass, 1995). Aphids which survived an attack had lower fecundity but equal longevity relative to aphids that were not attacked (van Steenis and El-Khawass, 1995).
There has been little study of the effect of mites on A. gossypii. Studies of Allothrombium pulvinum (Zhang et al., 1993; Zhang and Chen, 1993) are limited to the mite's distribution in the field.
The most important A. gossypii pathogens discovered to date are all fungi. All A. gossypii life stages may be attacked.
The two best-studied pathogens are Neozygites fresenii and Verticillium lecanii. Several other fungal pathogens have also been reported: Arthrobotrys sp., Erynia aphidis and Erynia delphacis (Shimazu, 1977; Sanchez-Peña, 1993).
N. fresenii takes 3, 4, 5-6 and 6-8 days to develop at temperatures of 30, 25, 20 and 15°C, respectively. Furthermore, at 35°C it did not kill aphids (Steinkraus et al., 1993). N. fresenii was able to produce up to 9835 conidia from a single aphid. The number of conidia was correlated with aphid size, but it has been suggested that handling or storage properties of larger aphids could explain this observation (Steinkraus et al., 1993).
N. fresenii can be a major cause of aphid mortality in cotton grown in the Texas/Arkansas area of the USA Steinkraus et al., 1991, 1993). It has a distinct diurnal periodicity in spore discharge, with greatest activity occurring between the hours of 01.00 and 05.00 with almost no activity between 09.00 and 24.00 h (Steinkraus et al., 1996). It has not been reared on artificial media, but Steinkraus et al. (1993) reported on propagation in an A. gossypii colony and the longevity of N. fresenii in cold storage.
V. lecanii is an important source of mortality for aphids under greenhouse conditions, but there are no reports of its impact on A. gossypii under field conditions. Its effectiveness is emphasized by its use as an aphicide in commercial greenhouses in the UK (Hall, 1985; Sopp et al., 1990). Its success in this capacity is partly due to the ability of V. lecanii to grow in artificial media. As one might expect, different strains of V. lecanii show different growth rates and different levels of pathogenicity (Hall, 1982; Kitazawa et al., 1984; Yokomi and Gottwald, 1988).
Temperatures above 35°C and humidity below 85% inhibit conidial discharge from A. gossypii cadavers (Steinkraus and Slaymaker, 1994). During the infection process, primary conidia germinate and form capilliconidia. This process is so sensitive to humidity that at 95% RH and 25°C, 90% of the primary conidia will germinate, whereas at 89% RH only 19% will germinate (Steinkraus and Slaymaker, 1994).
Means of Movement and DispersalTop of page
Long distance dispersal is by wind. Aerial sampling may be suitable for predicting future aphid abundance, but was not suitable for predicting existing populations (Parajulee et al., 2003). A. gossypii was collected at 150 m in India, and it was concluded that this dispersal of A. gossypii was over tens or hundreds of kilometres (Reynolds et al., 1999).
Plant TradeTop of page
|Plant parts liable to carry the pest in trade/transport||Pest stages||Borne internally||Borne externally||Visibility of pest or symptoms|
|Bark||arthropods/eggs||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Flowers/Inflorescences/Cones/Calyx||arthropods/adults; arthropods/nymphs||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Fruits (inc. pods)||arthropods/adults; arthropods/nymphs||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Leaves||arthropods/adults; arthropods/nymphs||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Seedlings/Micropropagated plants||arthropods/adults; arthropods/nymphs||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Stems (above ground)/Shoots/Trunks/Branches||arthropods/adults; arthropods/nymphs||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Plant parts not known to carry the pest in trade/transport|
|Growing medium accompanying plants|
|True seeds (inc. grain)|
Impact SummaryTop of page
|Fisheries / aquaculture||None|
ImpactTop of page
A. gossypii is extremely polyphagous and very damaging to many economically important crops, including cotton, aubergine, citrus, coffee, melon, okra, peppers, potato, squash and sesame. It is a major pest of cotton and cucurbits. A. gossypii has a worldwide distribution, although in arctic regions it is mostly confined to glasshouses. It is particularly abundant in the tropics.
Economic damage due to A. gossypii is by direct feeding, the excretion of honeydew and virus transmission. Damage to cotton, okra and certain cucurbits occurs when large populations of aphids build up, feed on the crops and excrete honeydew. However, its biggest overall economic impact is as a vector of pathogenic plant viruses in over two dozen crops. There is little quantitative information on exact crop losses. In cotton, for example, A. gossypii is only one of many crop pests. Monetary losses to this pest are substantial and are a result of crop loss and crop quality reduction, and the expense of pesticides.
A. gossypii causes direct feeding damage to cotton, okra and some other crops, by sucking the sap directly from the phloem, causing the removal of nutrients. The drain on plant nutritional resources can be considerable. Adverse physiological responses of plants to direct feeding can also occur. The undersides of young leaves are preferred, but the entire plant may be covered when populations are large. Infested leaves curl downwards and may appear wrinkled or reddened. Heavy infestations can result in wilting. Young plants often have reduced or stunted growth, and may sometimes be killed. A. gossypii is the principal aphid attacking cotton, on which it is an early through mid-season pest; although damaging late season infestations can occur, especially if broad-spectrum insecticides have reduced natural enemy populations (Matthews, 1989; Ebert and Cartwright, 1997).
In the USA, A. gossypii caused more insect-related damage to cotton than any other pest in 1991. Of 13 million acres harvested, around 10 million acres were classified as infested with aphids, resulting in losses of over 360,000 bales (Head, 1992). Losses in Texas alone were around 333,000 bales, representing a yield loss of approximately 6%. Yield reductions of over 100 pounds of lint per acre are not uncommon (Price et al., 1983). In a study of late season aphids, also in Texas, one part of a field was treated to keep aphid populations below 50/leaf, while the other part had aphid densities greater than 50/leaf. In 1992, aphid populations that exceeded 50/leaf for 3 weeks, and 100/leaf for 2 weeks significantly reduced cotton plant height. The higher aphid levels reduced lint yield by 16% in 1992 and 24% in 1993, with a 14% and 25% reduction in seed yield, respectively. None of the cotton quality parameters measured were affected. Differences in gross returns between low aphid plots and high aphid plots in the 2 years were $65.32 and $99.69 per acre (Fuchs and Minzenmayer, 1995). In the USA, the movement onto cotton from noncultivated host plants was an important factor determining A. gossypii infestation; this movement was gradual and ongoing from the time cotton seedlings emerged (O'Brien et al., 1993).
In California, USA, seed yield was reduced by 0.21 lbs seed per aphid-day for cotton planted early in the season (Godfrey and Wood, 1998). However, there was no yield loss in cotton planted later, even though aphid infestation levels were similar. Yield losses were due to a 13.8% decrease in the number of bolls, and a 5.7% decrease in boll size.
The impact of a pest complex on upland cotton in India was calculated in four regions. Aphids, mainly A. gossypii, were more numerous on plants in treatment, compared with control, plots in all cases. Insecticides (soil treatment and regular spraying) resulted in yield (kg/ha) increases between 58.4 and 75.1%. In economic terms, taking into account the cost of plant protection and the net income from the cotton, an additional income of between 4340 and 6115 Rupees/ha was obtained from the pest control treatments (Sivaprakasam and Balasubramanian, 1981). In another Indian study, yield losses in cotton due to sucking pests were between 20.90 and 26.30%, with a glabrous hybrid giving consistently higher yields than a hairy hybrid (Kulkarni and Raodeo, 1986). Reductions in cotton yield due to sucking pests by 16.2 to 55.6% have been reported in studies from Russia and Brazil (Moskovetz, 1941; Vendramin and Nakano, 1981). In cotton in Zambia, A. gossypii damage caused up to 80% yield loss, while a survey of that country's farmers ranked it as the most serious cotton pest (Javaid et al., 1987).
In China, A. gossypii infestations are most serious in the seedling stage, particularly in the northern cotton zone. In one study, cotton seedlings (3 leaves or less) were more sensitive to infestations, while damage on older plants was lower, in part due to compensatory growth effects when precipitation was sufficient. Seedlings were stunted, with a decreased leaf area index and reduced root system development, while the time to squaring was delayed. A damage index, used to establish thresholds for spraying, resulted in adequate control with a 50% reduction in insecticides (Zhang et al., 1982). In Australia, a prototype IPM system maintained cotton yields, while reducing insecticide usage by around 40 to 50% (Hearn et al., 1981).
In Spain, A gossypii is capable of causing 55% yield loss in clementines (Citrus clementina) (Mendoza et al., 2001). Mendoza et al. (2001) provided an estimate of the economic injury level (EIL) and the environmental EIL. The EIL for A. gossypii was 271 aphids/m²; however, this figure will change with the cost of control, the sale price, and yield loss due to the aphid change.
Direct feeding damage by A. gossypii on cotton is related to plant growth stage and level of aphid infestation. Aphid populations increase rapidly with favourable climatic conditions and plant nutritional quality. Levels of damage are influenced by the presence of natural enemies and biological control, pesticide efficacy, the presence of pesticide resistance in aphids, and compensatory growth in plants (Zhang et al., 1982; Slosser et al., 1989). The planting date was the most important variable, in a multiple regression analysis, affecting aphid density in dryland cotton in Texas, USA (Slosser et al., 1989). Optimum temperature for population growth is around 20-25°C (Akey and Butler, 1993). Light intensity and daylength significantly influence rate of population increase, while heavy rain can directly reduce populations by washing them off leaves (Ebert and Cartwright, 1997).
Natural enemies are important in many areas in preventing secondary pest outbreaks. Reduction of natural enemies by insecticides therefore might exacerbate aphid attack by removing this natural control. In a Chinese study, for example, chemical control of the pest complex during the early season led to increased damage to cotton by A. gossypii during the mid and late season (Chen et al., 1991). In predator exclusion experiments in the USA, large decreases in cotton lint yield were observed in caged plots compared with uncaged plots where natural enemies could control aphid numbers. Fibre quality was also reduced by high aphid numbers in predator exclusion cages (Kidd and Rummell, 1997). The reduction in natural enemies resulting in greater aphid abundance was also found in Egyptian cotton fields (Abou-Elhagag, 1998).
The relationship between infestation, nitrogen application and cotton yield was investigated in a study in the Philippines. The more nitrogen applied, the greater was the infestation by A. gossypii, but this was outweighed by the increased yields obtained due to nitrogen fertilizer applications (Villamayor, 1976).
In okra, A. gossypii, along with a jassid bug (Amrasca biguttula biguttula), reduced total yield by 19% in a study in Bangalore, India (Srinivasan and Krishnakumar, 1983). A. gossypii causes most damage at the seedling stage in okra, with feeding reducing the vigour of plants (Pareek et al., 1987).
Losses due to honeydew, excreted by feeding A. gossypii, can be considerable. Honeydew interferes with leaf transpiration, and acts as a substrate for fungi, including sooty moulds (Capnodium), which blacken leaves and reduce photosynthetic efficiency. Honeydew can also act as an attractant to other crop pests, and insects such as bees, wasps and ants, that may provide protection for the aphids from their natural enemies (Slosser et al., 1989). The presence of honeydew contaminates cotton lint, reducing its quality and economic value (Slosser et al., 1989). The presence of honeydew on fruit crops can significantly reduce their marketability.
A. gossypii transmits over 50 plant viruses, including non-persistent viruses of beans and peas, crucifers, celery, cowpea, cucurbits, Dahlia, lettuce, onion, papaya, peppers, soyabeans, strawberry, sweet potato, tobacco and tulips (Blackman and Eastop, 2000). In cotton, it transmits Cotton anthocyanosis virus, Cotton curliness virus, cotton blue disease, Cotton leaf roll and purple wilt (Kennedy et al., 1962; Brown 1992). In addition to Cotton anthocyanosis, Lily rosette, Lily symptomless and Pea enation mosaic virus, are all transmitted in a persistent manner (Blackman and Eastop, 2000). On the majority of crops it attacks, the ability of A. gossypii to act as a vector of plant viruses can result in significant economic losses (Ebert and Cartwright, 1997).
A. gossypii is an important vector of Papaya ringspot virus, transmitting both the P (PRSV-P) and W (PRSV-W) strains. The former is a disease of papaya, whereas PRSV-W, also called Watermelon mosaic virus 1 (WMV-1), infects cucurbits and watermelon (Kessing and Mau, 2000). A. gossypii also transmits Watermelon mosaic virus 2 (WMV-2), Zucchini yellow mosaic virus (ZYMV) and Celery mosaic virus (CeMV). These potyviruses are transmitted in a non-persistent manner.
A. gossypii is the most important vector of Cucumber mosaic virus (CMV) in cucurbits. CMV has one of the widest host ranges of any plant virus. It can be acquired in 5-10 seconds and be transmitted in less than 1 minute. The ability of CMV to be transmitted declines after about 2 minutes and is usually lost within 2 hours (Francki et al., 1985). Citrus tristeza virus (CTV) is transmitted in a non- or semi-persistent manner by A. gossypii; the virus remains infectious for 24 hours, and is specific to plants in the Rutaceae (Brunt et al., 1996).
A. gossypii is the only known vector of the virus causing cotton blue disease in Africa. In field trials in the Central African Republic, yield losses due to A. gossypii and cotton blue disease were significantly reduced with insecticide treatments. A combination of organophosphorus and pyrethroid insecticides was recommended for control (Cauquil et al., 1978; Cauquil, 1981). In Chad, all cotton varieties were found to be susceptible to cotton blue disease, except one with a small degree of tolerance. Cotton blue disease decreased the length of fibres by about 2.5%, compared with uninfested controls, although other quality characteristics were unaffected (Dyck, 1979).
Environmental ImpactTop of page
Impact: BiodiversityTop of page
Social ImpactTop of page
Detection and InspectionTop of page
In addition to looking for symptoms on the host plant (chlorosis, distortion) one should look for the presence of sticky surfaces indicative of aphid feeding on upper leaves, and the sooty moulds which often grow on the honeydew.
A. gossypii is often attended by ants. A small group of ants walking on the foliage may often be associated with the presence of aphids. All of these symptoms are general indications of homopteran (aphid, whitefly, mealybug, etc.) feeding and are not unique to A. gossypii.
Position on Plants
On aubergines in India, A. gossypii settled on older mature leaves. It moved to younger tissues only when population pressure forced it to; thus aphid populations were always greatest on the older leaves (Banerjee and Raychaudhuri, 1985; Ebert and Cartwright, 1997).
On cantaloupes (Cucumis melo) in the USA, A. gossypii was most abundant on the basal portion of vines (Edelson, 1986; Ebert and Cartwright, 1997).
On cotton in the USA, it was most abundant in the middle canopy, followed by the upper canopy. However, this pattern may be a result of high aphid mortality from a fungal pathogen in the lower canopy rather than a result of aphid behaviour (O'Brien et al., 1993; Ebert and Cartwright, 1997).
In general, A. gossypii is not usually a post-harvest pest. However, damage done in the field can cause post-harvest problems. Such damage frequently occurs due to contamination with honeydew. Prevention can only occur prior to harvest.
One possible situation where A. gossypii could be a post-harvest pest is in the cut flowers industry. The symptoms to look for would be identical to those for the whole plant.
Similarities to Other Species/ConditionsTop of page
Taxonomically A. gossypii is difficult to separate from Aphis frangulae.
Prevention and ControlTop of page
Due to the variable regulations around (de)registration of pesticides, your national list of registered pesticides or relevant authority should be consulted to determine which products are legally allowed for use in your country when considering chemical control. Pesticides should always be used in a lawful manner, consistent with the product's label.
Despite a number of experimental studies on the natural enemies of A. gossypii (see Natural Enemies), there are few records available of biological control being practised in the field. One successful study using biological control was in Egypt. Two field releases of Chrysoperla carnea at a ratio of 1:5 (predator:aphid) eliminated the aphid in 12 days, whereas it took a single release of Coccinella undecimpunctata at a ratio of 1:50 to get 99.7% control in okra (Zaki et al., 1999).
Potts and Gunadi (1991) reported a decrease in A. gossypii populations in potatoes that are intercropped with Allium cepa or Allium sativum. To achieve the reduction, the onions had to be planted within 0.75 m of potato plants. However, intercropping poses a problem when the minor crop harbours a disease of the primary crop. Such a system has been documented in Taiwan where bananas were interplanted with cucumbers (an alternative host for Banana mosaic virus) (Tsai et al., 1986). A similar effect also occurs when alternative hosts (of aphid and virus) are in neighbouring fields (Tsai et al., 1986).
In cotton, an unusual approach was to top the plants after boll opening. This removed the top leaves where aphids fed, and thereby reduced contamination of bolls below these leaves. Topping was done by hand, using a pruning knife to remove the terminal spray of each plant (Deguine et al., 2000).
Glabrate cotton supported fewer aphids than more pubescent cotton (Dunnam and Clark, 1938; Weathersbee et al., 1994). However, pubescence has the opposite effect on A. gossypii feeding on muskmelons (Kennedy et al., 1978; Ebert and Cartwright, 1997). Greater trichome density resulted in fewer aphids and less virus disease in ashgourd, Benincasa hispida (Khan et al., 2000).
Many crops have some level of physiological resistance to A. gossypii that can be classified into one of three categories: tolerance, antixenosis or antibiosis. The causes for resistance were examined extensively in muskmelons and cucumbers (Ebert and Cartwright, 1997). Resistance has also been documented in okra (Uthamasamy et al., 1976; Gunathilagaraj et al., 1977); Gossypium hirsutum and Gossypium arboreum (Chakravarthy and Sidhu, 1986) (Reed et al., 1999); Antigastra catalunalis (Muralidharan et al., 1977); Citrullus lanatus (MacCarter and Habeck, 1973); Solanum melongena (Sambandam and Chelliah, 1970); Dendranthema morifolium x D. indicum (Storer and van Emden 1995) and Colocasia esculenta (Palaniswami et al., 1980).
While host plant resistance is easily classified into three categories, the cause-effect relationships are not often clear because studies focus on one part (toxicity) and ignore other aspects that may be correlated (genetically linked, or correlated within the study plants but without a genetic basis). Susceptibility to A. gossypii was attributed to high protein and high amino acid content of cucumber cultivars (Ahmed, 1994). Differences in trichome density and differences in toxins were not measured. Likewise, in studies of host-plant resistance, nutritional suitability was not usually measured.
In melons, resistance is conferred by the Vat gene. The cause of the resistance appears to be due to a modified phloem sealing physiology that reduces the quantity of sap an aphid can extract from each feeding site. Furthermore, the phloem they do get has reduced total protein (Chen et al., 1997).
A novel approach to pest management has been the idea of eliciting natural defences in plants using mechanical wounding, infection, or sprays of elicitors. Jasmonic acid applied to cotton plants in California, USA, reduced survival and decreased the number of progeny per leaf (Omer et al., 2001). A similar experiment was done in China, where cotton plants were physically wounded, and some wounds were infected with a bacterium (Pseudomona gladioli D-2251). Wounded plants had fewer aphids and it was less likely that wounded plants would be infested. Infestation frequency and aphid abundance on infested plants was further reduced if the bacterium was present (Li et al., 1998). The phenomenon is widespread, and occurs naturally. A similar study was conducted on cucumber with infection by Cladosporium cucumerinum (Moran, 1998), but without physically damaging the plants. In this system A. gossypii was unaffected by the presence or absence of the fungus, whereas damage from another fungal pathogen was reduced. Wind induced mechanical stress can increase peroxidase activity in cucumber, and with at least 12 days of wind stress can result in detectable reductions in aphid populations (Moran and Cipollini, 1999). However, Moran also found that this mechanical stress can increase pathogen susceptibility.
The opposite of host plant defence would be induced susceptibility. For example, cotton plants fertilized with high nitrogen are better hosts for A. gossypii, and this results in greater damage (Cisneros and Godfrey, 2001; Nevo and Moshe, 2001).
A wide array of chemicals have been used to control A. gossypii but there is a persistent problem with resistance development. In a few cases, chemical treatments have resulted in more damage than would have occurred without the treatment. The mechanism for this effect is not known, but contributing factors include the removal of beneficial organisms (predators and parasites), stimulation of aphid reproduction, and changes in plant physiology due to the application of the chemical.
Resistance development has been a problem associated with the following insecticides.
Acephate (Whalon, 2003)
Alpha-cypermethrin (Whalon, 2003)
Bifenthrin (O'Brien et al., 1992); 19 times more resistant at LD50 in Australia (Herron et al., 2000)
Chlorpyrifos (O'Brien et al., 1992)
Carbaryl: 31 times more resistant at LD50 in Japan (Saito and Hama, 2000)
Chlorpyrifos (Whalon, 2003)
Cyhalothrin (Whalon, 2003)
Cypermethrin (Whalon, 2003)
Deltamethrin (Gubran et al., 1992); 19 times more resistant at LD50 in Australia (Herron et al., 2001)
Demeton (Whalon, 2003)
Diazinon (Furk and Hines, 1993; Saito et al., 1995)
Dimethoate (Gubran et al., 1992); in China, Greece, Peru, UK, Zimbabwe (Han et al., 1998)
Endrin (Whalon, 2003)
Esfenvalerate (Whalon, 2003)
Fenitrothon: (Saito et al., 1995); 16 times more resistant at LD50 in Japan (Saito and Hama, 2000)
Fenpropathrin (Whalon, 2003)
Fenvalerate (Gubran et al., 1992); 29,000 times more resistant at LD50 in China (Wang et al., 2002)
Imidacloprid: 8 times more resistant at LD50 in China (Wang et al., 2002)
Lambda-cyhalothrin: 28 times more resistant at LD50 in Australia (Herron et al., 2000)
Malathion (Saito et al., 1995); 27 times more resistant at LD50 in Japan (Saito and Hama, 2000)
Oxydeprofos (Whalon, 2003)
Permethrin: in China, Greece, UK (Han et al., 1998)
Phoxim: in China (Xie et al., 2002)
Pirimcarb (Gubran et al., 1992; Furk and Hines, 1993; Silver et al., 1995; Han et al., 1998; Rongai et al., 1998); 1700 times more resistant at LD50 in Australia (Herron et al., 2001); in China, Greece, Peru, UK, Zimbabwe (Han et al., 1998); Italy (insensitive at all concentrations tested) (Rongai et al., 1998)
Profenofos: 6 times more resistant at LD50 in Australia (Herron et al., 2001)
Sulprophos (Kerns and Gaylor, 1993)
y-BHC (Gubran et al., 1992)
Aphids reared on different plants show different levels of susceptibility to insecticides. A. gossypii has manifested several mechanisms for resistance: enzymatic differences, target insensitivity, and modifications to the cuticle. However, studies in this area suffer from an inability to differentiate between phenotypic plasticity versus differences in aphid genotype as selected by host plant (Ebert and Cartwright, 1997).
Resistance to fenvalerate was increased 29,000 after 16 generations of selection in a strain reared on cotton or cucumber, and resistance to imidacloprid was increased 8.1 fold in 12 generations in a greenhouse environment (Wang et al., 2002). There was cross resistance to fenvalerate in aphids exposed to imidacloprid, but aphids exposed to fenvalerate did not show a cross resistance to imidacloprid. Resistance to fenvalerate was greater in aphids on cotton, but there did not appear to be a host plant difference in resistance levels to imidacloprid (Wang et al., 2002).
Host-plant-mediated susceptibility for endrin, malathion, ethyl parathion, dimethoate and carbaryl was reported for A. gossypii reared on six cucurbitaceous hosts (Juneja and Sharma, 1973). A. gossypii on cucumbers were consistently less susceptible to all insecticides relative to aphids from the original culture on bottle gourds (Lagenaria siceraria). Other than this, there was no consistency in the level of resistance and host plant.
Toxicity rank of pesticides changed depending on host plant (Juneja and Sharma, 1973). A. gossypii colonies feeding on watermelons (in comparison with cotton) showed differences in pesticide susceptibility between bifenthrin, chlorpyrifos and dimethoate (McKenzie and Cartwright, 1994).
There can be an interaction between host-plant resistance and pesticide resistance in A. gossypii. In some cases, pesticide resistance in A. gossypii was correlated with different aliesterase levels in aphids on different host plants. Aphids reared on melons or cucumbers showed elevated aliesterase levels relative to aphids from aubergines or potatoes (Saito, 1991). A. gossypii with high aliesterase activity maintained original levels of aliesterase activity even when moved to solanaceous crops, and aphids with low aliesterase activity maintained low levels when moved to cucurbitaceous crops. This represents a difference between two strains of A. gossypii correlated with a difference in host plant, and is clearly not host induced.
Significant differences in esterase patterns and esterase quantity have been reported, and differences with insecticide resistance (Furk et al., 1980; Hama and Hosoda, 1988; Takada and Murakami, 1988; O'Brien, 1992; Saito et al., 1995) and host-plant preference (Furk et al., 1980) have been correlated. While mixed function oxidases may play a role in detoxification reactions, the esterases and carboxylesterases showed more conspicuous differences between susceptible and resistant aphid strains (Sun et al., 1987).
The biological cost of resistance for an organophosphate-resistant aphid feeding on cotton has been examined. Over the first few days resistant alate A. gossypii produced more progeny, but over the life span of the adult there was no significant difference. However, the reproductive rate during the first few reproductive events are the most influential in determining population growth rate (O'Brien, 1992).
Peroxidase levels in salivary glands, sheath material and salivary excretions of A. gossypii might play some role in detoxifying systemic insecticides. For this reason, the role of salivary gland enzymes in detoxification has been examined as a possible defence. While the peroxidases were effective in detoxifying hordenine and gossypol, the role of these enzymes in the natural habitat of A. gossypii is not clear (Miles and Peng, 1989).
A. gossypii also acquires resistance through target site insensitivity and through increased production of the affected enzymes. Acetylcholinesterase (AChE) insensitivity was shown in A. gossypii resistant to organophosphate and carbamate insecticides (Moores et al., 1988). Furthermore, activity level of AChE has been correlated with pirimicarb resistance (Suzuki and Hama, 1994; Silver et al., 1995). For two clones this resistance was shown to be specific to pirimicarb (Silver et al., 1995). These clones were 800 times more resistant than the susceptible clone, but no more than 22 times more resistant to six organophosphates, six carbamates and two pyrethroids. The mechanism was linked to AChE with higher catalytic activity and lower affinity to pirimicarb in the resistant clones. Cuticular differences as well as AChE insensitivity played a role in insecticide resistance in paraoxon (Sun et al., 1987).
Life stage is another factor affecting the susceptibility of A. gossypii to insecticides. Alate adults on cotton were more resistant than apterous adults to chlorpyrifos and biphenate (Grafton-Cardwell, 1991).
Factors which may result in pest resurgence (other than a reduction in natural enemies) have been examined. Cotton fields treated with sulprophos had elevated numbers of A. gossypii, but the cotton plants in these fields had significantly elevated levels of threonine and essential amino acids (Kerns and Gaylor, 1993). Cotton plants treated with dimethoate had larger aphids, but treated plants had lower sugar content, lower nitrogen content, lower carbohydrate to nitrogen ratio, and higher amino acid content (Sithanantham et al., 1973).
Although altered plant physiology may account for A. gossypii resurgence in some cases, it is not the only cause. Direct applications of deltamethrin and carbaryl stimulated the reproductive rate of A. gossypii. Some of this effect was due to elevated feeding levels at certain doses of deltamethrin (Gajendran et al., 1986).
Different host plants can play a direct role in detoxification of the insecticide (Owusu et al., 1995). Elevated A. gossypii populations have been reported following pesticide applications but the causes were not examined (Patel et al., 1986; Surulivelu and Sundaramurthy, 1986; Thimmaiah and Kadapa, 1986; Ebert and Cartwright, 1997).
Sub-lethal effects: Imidacloprid was observed to stimulate the production of alates in the USA (Conway et al., 2003). While six times as many alates were produced in imidacloprid treated plots, these aphids had significantly reduced fecundity.
Integrated Pest Management
Major IPM programmes have been developed for A. gossypii in cotton. There are computer-based systems under development in Australia (Hearn et al., 1981), Sudan (Abdelrahman and Munir, 1989; Stam et al., 1994), USA (O'Brien, 1992), Zambia (Javaid, 1993) and other countries (Silvie et al., 1993; Deguine et al., 1994).
Transgenic plants are used to manage other pests in crops where A. gossypii occurs. In China, it was shown that A. gossypii develops resistance to Bt, and Bt+CpTI transgenic cottons, and after only three generations, the aphids had longer reproductive duration, greater survival, and greater maximum potential fecundity than similar aphids on non-transgenic cotton cultivars (Liu et al., 2005).
Economic Threshold Levels
Despite the lack of quantitative data on exact yield reductions caused by A. gossypii, there are a number of reports on the control action threshold. In Sudan on cotton, this level was 30% infestation of the plants during the first 2 months of the season (Stam et al., 1994). Using a damage index, a rating of 250 for seedling cotton with fewer than three leaves, and a rating of 350-400 for larger plants was recommended for treating cotton in China (Zhang et al., 1982).
In Spain, the economic injury level (EIL) in clementines (Citrus clementina) and the environmental EIL was estimated. The EIL for A. gossypii was 271 aphids/m²; however, this figure will change with the cost of control, the sale price, and yield loss due to the aphid change (Mendoza et al., 2001).
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